Irisin May Boost a Key Protein to Help Muscle Loss and Energy Problems Caused by Stress Hormones

Male C57BL/6J mice (8 weeks old) were housed under controlled conditions (temperature: 22 ± 3 °C; humidity: 40–50%; 12 h light-dark cycle) with ad libitum access to food and water. Animals were randomly assigned to five groups (n = 6 per group) and administered treatments as follows: (i) Control group: vehicle (PEG300 solution, i.p.); (ii) Irisin group: The irisin stock solution was prepared using sterile physiological saline as the solvent at a concentration of 500 μg/mL (2.5 mg/kg, i.p.); (iii) Dex group: The formulation of the dexamethasone stock solution comprises PEG300 at a concentration of 2.5 mg/mL (25 mg/kg, i.p.); (iv) Dex + Irisin group: dexamethasone (25 mg/kg, i.p.) + irisin (2.5 mg/kg, i.p.); (v) Dex + Irisin + EX-527 (SIRT1 inhibiting drug) group: dexamethasone (25 mg/kg, i.p.) + Irisin (2.5 mg/kg, i.p.) + EX-527 (10 mg/kg, i.p.). Based on previous studies [16], Irisin was administered three times weekly, while dexamethasone and EX-527 were administered daily, over a total duration of 14 consecutive days. Body weight was recorded daily. Samples of blood, gastrocnemius (GAS), and tibialis anterior (TA) muscles were collected at 24 h following the last injection. Upon collection, the muscle tissues were immediately weighed and photographed, followed by fixation in 4% paraformaldehyde or flash-freezing in liquid nitrogen for subsequent analysis. All experimental manipulations were carried out in compliance with protocols authorized by the Animal Care Committee of Shanxi Agricultural University (SXAU-EAW2024M.ID.011005320).

The mouse C2C12 myoblast cell line was acquired from the American Type Tissue Culture Collection (ATCC). Cells were maintained in high-glucose Dulbecco’s Modified Eagle Medium (BI, Beit HaEmek, Israel) supplemented with 10% fetal bovine serum (Sciencell, Carlsbad, CA, USA) at 37 °C in a humidified 5% CO₂ atmosphere. Upon reaching 70% confluency, the growth medium was replaced with a differentiation medium consisting of DMEM and 2% horse serum to induce myotube formation. Cultures were maintained under this condition for 4 days, with medium replacement daily, to induce multinucleated myotube formation. To induce myotube atrophy, fully differentiated myotubes were treated with 50 μM Dex (MCE, Monmouth Junction, NJ, USA) for 24 h. R-irisin (Phoenix Pharmaceuticals, Burlingame, CA, USA) and EX-527 (MCE, Monmouth Junction, NJ, USA) were administered concomitantly with Dex during the atrophy induction phase.

C2C12 myotubes, fully differentiated following seeding at an initial density of 1 × 10⁴ cells per well in 96-well plates, were subjected to a 24 h treatment with 50 μM dexamethasone (Dex) and varying concentrations of R-irisin (250, 500, 750, and 1000 ng/mL). Cell viability was subsequently assessed using a Cell Counting Kit-8 (CCK-8; YEASEN, Shanghai, China). Briefly, 10 μL of CCK-8 reagent was added to each well, followed by a 2 h incubation at 37 °C. The absorbance was then measured at 450 nm using a microplate reader (Molecular Devices, San Jose, CA, USA). The percentage of cell viability was calculated relative to the untreated control group using the formula: Cell Viability (%) = (As − A0)/(Ac − A0) × 100%, where As, A0, and Ac were the absorptions of test sample, back-ground, and control (DMSO), respectively.

Following fixation with 4% paraformaldehyde (15–20 min) and permeabilization with 0.1% Triton X-100 (10 min), the cells were blocked for 30 min at room temperature using 5% goat serum (Solarbio, Beijing, China). Subsequently, the cells were incubated overnight at 4 °C with a primary antibody targeting myosin heavy chain (MyHC). After extensive washing, a species-matched Alexa Fluor-conjugated secondary antibody was applied for 1 h at 37 °C. Nuclei were visualized by counterstaining with DAPI for 10 min at room temperature. Fluorescence images were acquired with a Eclipse Ts2R microscope (Nikon, Yokohama, Japan) and processed using the NIS-Elements software (v4.20).

Muscle tissues were fixed in 4% paraformaldehyde for 24 h, followed by graded ethanol dehydration and xylene clearing. Paraffin-embedded tissues were sectioned into 0.8 μm-thick slices using an automated rotary microtome (LEICA, Hesse, Germany). The sections were deparaffinized with xylene, rehydrated through graded ethanol series, and stained with haematoxylin and eosin. After mounting with neutral balsam (Solarbio, Beijing, China) images were captured and processed using a microscope (Nikon, Yokohama, Japan) equipped with NIS-Elements imaging software.

Protein extracts were prepared from muscle tissues or myotubes by homogenization in cold RIPA buffer (Beyotime Biotechnology, Shanghai, China) containing a cocktail of protease and phosphatase inhibitors. Following lysis on ice for 30 min and centrifugation at 13,000× g for 5 min at 4 °C, the supernatant was collected. Protein concentration was determined using a BCA assay kit (Beyotime, Shanghai, China). Equal amounts of protein were then resolved by SDS-PAGE and electrophoretically transferred onto PVDF membranes (Millipore, Burlington, MA, USA). The membranes were blocked with 5% skimmed milk in TBST for 1 h at room temperature before being incubated with primary antibodies overnight at 4 °C. After washing, the membranes were probed with appropriate HRP-conjugated secondary antibodies for 1 h at 37 °C. Signal detection was performed using enhanced chemiluminescence reagents (ABBKINE, Wuhan, China), and band intensities were captured with a ChemiDoc XRS+ imaging system (Bio-Rad, Hercules, CA, USA). Quantitative analysis was conducted using ImageJ software (v1.47, National Institutes of Health, Bethesda, MD, USA) after normalizing the band intensities to the endogenous control GAPDH.

To distinguish the acetylated form of PGC-1α from its total protein levels, we conducted immunoprecipitation (IP) followed by Western blotting. In this procedure, cell lysates were incubated with anti-PGC-1α antibody and A/G agarose beads (Santa Cruz Biotechnology, Santa Cruz, CA, USA) overnight at 4 °C under constant rotation. The beads were then pelleted and washed extensively with cold RIPA buffer. Bound proteins were eluted by heating in 5×SDS sample buffer and subjected to SDS-PAGE. Immunoblotting was subsequently carried out, employing an antibody against acetylated lysine to detect PGC-1α acetylation, with total PGC-1α serving as the internal control.

C2C12 myoblasts were seeded in confocal culture dishes and allowed to reach full differentiation. Differentiated myotubes were with dexamethasone (50 μM), R-irisin (1000 ng/mL), and the SIRT1 inhibitor EX-527 (20 μM) for 24 h. According to the manufacturer’s instructions, live cells were stained with JC-1 fluorescent dye (Beyotime Biotechnology, C2003S, China) at 37 °C for 20 min to visualize the level of mitochondrial membrane potential. Mitochondrial membrane potential was visualized by capturing fluorescence images using a confocal laser scanning microscopy (CLSM) system (Olympus, Yokohama, Japan).

ATP concentrations were measured following the instructions of a commercial detection kit (Beyotime Biotechnology, S0026, China). Muscle samples were homogenized in ice-cold lysis buffer and centrifuged (12,000× g, 5 min, 4 °C). The supernatant was then incubated with the ATP detection reagent, and luminescence was recorded on a FilterMax F5 microplate reader (Molecular Devices, San Jose, CA, USA). Sample ATP levels were interpolated from a standard curve.

SIRT1 deacetylase activity was quantified using a fluorometric assay kit (Elabscience^®, Wuhan, China) following the manufacturer’s instructions. Briefly, 1 × 10⁶ cells were homogenized in 0.2 mL physiological saline, centrifuged at 12,000× g for 5 min at 4 °C, and the supernatant was collected. The cell lysate was incubated with reaction buffer in a 96-well plate, and fluorescence intensity was monitored at excitation/emission wavelengths of 340/440 nm using a microplate reader (Molecular Devices, San Jose, CA, USA). Enzyme activity was normalized to total protein content.

Intracellular ROS levels were assessed using a commercially available ROS detection assay kit (Beyotime Biotechnology, Shanghai, China). C2C12 myoblasts were seeded in sterile cell culture dishes and allowed to reach full differentiation. Differentiated myotubes were incubated with the fluorescent probe DCFH-DA (dichlorofluorescin diacetate) at 37 °C for 20 min. Fluorescence intensity was quantified using a fluorescence microscope (Nikon, Yokohama, Japan) equipped with appropriate excitation/emission filters (Ex/Em = 488/525 nm).

The concentration of irisin in plasma and cell supernatants from each mouse group was quantified using a species-specific enzyme-linked immunosorbent assay (ELISA) kit (Elabscience^®, Wuhan, China) in strict accordance with the manufacturer’s instructions.

Exhaustion Time and Distance. Following a 2-day acclimation period, running performance was assessed as previously described [18], Mice ran on a treadmill (SANS Biotechnology, Nanjing, China) with a 13% incline. The speed was increased by 5 cm/s every 2 min until exhaustion, which was designated when a mouse’s hindlimbs remained on the electric grid for >10 s. The total time to exhaustion and running distance were recorded.

Limb Grip Strength. Forelimb and hindlimb strength was measured using a dynamometer (SANS Biotechnology, Nanjing, China). Mice were allowed to grasp a horizontal grid and were pulled backward ten times. The peak force (in Newtons) applied before the mouse lost its grip was recorded for each trial. The average of ten measurements was normalized to the animal’s body weight.

Inverted Grid Test. Motor coordination and endurance were evaluated by placing mice on a wire grid, which was then inverted over a 2 s period. The grid was held 40–50 cm above a soft surface. The latency to fall was recorded as the hanging time.

The SUnSET assay was employed to assess the global protein synthesis rate. Briefly, puromycin (1 μg/mL) was administered to the culture medium 10 min before cell harvesting. The resulting cell extracts were then subjected to Western blot analysis using an anti-puromycin antibody (12D10).

Ultrastructural analysis of muscle tissue was performed by transmission electron microscopy (FEI, Hillsboro, OR, USA). Briefly, samples were fixed in 1% glutaraldehyde, sectioned, and mounted on copper grids for observation under the microscope equipped with a high-resolution camera.

Cell apoptosis was assessed with an Annexin V-FITC/PI detection kit (ABBKINE, Wuhan, China) per the manufacturer’s protocol. In brief, 1 × 10⁵ cells per sample were resuspended in binding buffer and stained with Annexin V-FITC and propidium iodide (PI) for 15 min at room temperature in the dark. Annexin V is a calcium-dependent phospholipid-binding protein that specifically recognizes and binds to phosphatidylserine (PS) externally exposed on the cell membrane. PI, a nucleic acid-binding dye, enters the cell and binds to nuclear DNA during the late stage of apoptosis when the cell membrane integrity is disrupted. The samples were then immediately analyzed on a BD LSRFortessa™ flow cytometer (Becton, Dickinson and Company, Franklin Lakes, NJ, USA), configuring the FITC (Ex/Em: 488/520 nm) and PI (Ex/Em: 535/615 nm) channels. The resulting data were processed using FlowJo software (v10.8.1, BD Biosciences, Franklin Lakes, NJ, USA).

All the statistical data are presented as the mean ± standard error of the mean (SEM). Comparisons between two groups were performed using a two-tailed Student’s t-test, and comparisons among multiple groups were performed using one-way analysis of variance (ANOVA) with Tukey’s post hoc test. A p value of less than 0.05 (p < 0.05) was considered to indicate statistical significance. All the data were drawn using GraphPad Prism 8.3.

To investigate the role of FNDC5/irisin in glucocorticoid-induced muscle atrophy, we employed Dex, a synthetic glucocorticoid analogue, to establish a well-characterized murine model of muscle wasting. Dex administration caused a marked reduction in body weight over the treatment period (Figure 1A). Consistent with systemic wasting, Dex treatment resulted in significant decreases in gastrocnemius and tibialis anterior muscle weights (Figure 1B) and reduced myofibre cross-sectional area in the gastrocnemius (Figure 1C). Notably, Western blot analysis confirmed elevated expression of atrophy-related ubiquitin ligases Atrogin-1 and MuRF-1 in Dex-treated muscles (Figure 1D,E). These data confirm that we successfully reproduced previously reported models of dexamethasone-induced muscle atrophy [19,20].

In parallel, FNDC5 protein levels in the gastrocnemius were significantly decreased following Dex exposure (Figure 1D,F). Strikingly, circulating irisin levels in plasma were also markedly reduced (Figure 1G). To translate these findings in vitro, we differentiated C2C12 myoblasts into myotubes and treated them with Dex. This treatment promoted myotube atrophy, manifested by decreased myotube diameter/fusion index (Figure 1H–J) and increased Atrogin-1/MuRF-1 expression (Figure 1L,M). Consistent with in vivo data, FNDC5 protein levels in Dex-treated myotubes were significantly diminished within 24 h (Figure 1N), accompanied by a sharp decline in secreted irisin levels in culture supernatants (Figure 1K). These observations suggest that suppression of the FNDC5/irisin axis may contribute to glucocorticoid-driven muscle wasting.

Given the observed downregulation of FNDC5/irisin in the Dex-induced muscle atrophy model, we investigated whether irisin reconstitution could rescue Dex-mediated myopathic phenotypes. In C2C12 myotubes, R-irisin administration dose-dependently reversed Dex-induced reductions in myotube viability (Figure 2A). Similarly, R-irisin treatment effectively prevented Dex-induced myotube atrophy (Figure 2B,C). Furthermore, even at the baseline level, the myotube diameter and fusion index were significantly increased following R-irisin exposure (Figure 2B–D). Western blot analyses revealed that R-irisin administration suppressed Dex-induced upregulation of atrophy markers Atrogin-1 and MuRF-1 (Figure 2E). Crucially, R-irisin robustly restored myosin heavy chain (MyHC) protein levels—a hallmark of myotube formation—in Dex-treated myotubes in a dose-dependent manner (Figure 2E). Strikingly, R-irisin (1 μg/mL) significantly enhanced baseline MyHC expression in control myotubes (Figure 2E), suggesting intrinsic myogenic properties. Furthermore, R-irisin demonstrated potent anti-apoptotic effects, markedly attenuating Dex-induced apoptosis in myotubes (Figure 2F,G). These findings collectively indicate that R-irisin functions as a pro-myogenic factor, promoting myotube hypertrophy while rescuing Dex-mediated atrophy and apoptotic responses.

To further validate the therapeutic efficacy of R-irisin in muscle wasting, we employed a robust glucocorticoid-induced atrophy model using dexamethasone. R-irisin treatment resulted in a significant elevation of circulating irisin levels in both control and dexamethasone-treated mice (Figure 3A). Moreover, a marked attenuation of dexamethasone-induced body weight loss was observed (Figure 3B). Consistent with systemic protection, R-irisin treatment markedly reduced Dex-mediated gastrocnemius and tibialis anterior muscle mass loss (Figure 3C,D). Histological analysis via haematoxylin and eosin (H&E) staining revealed that R-irisin preserved myofibre cross-sectional area in Dex-treated gastrocnemius muscles (Figure 3E,F). Notably, R-irisin treatment also induced a shift towards larger myofibre diameters in control mice (Figure 3E,F), suggesting pro-myogenic effects independent of atrophy reversal. Furthermore, R-irisin therapy improved functional outcomes, evidenced by enhanced grip strength, grid hanging performance, and treadmill exhaustive distance (Figure 3G–I). Collectively, these findings demonstrate that R-irisin exerts potent therapeutic efficacy against Dex-induced muscle atrophy and weakness.

To delineate the molecular mechanisms underlying R-irisin’s anti-atrophic effects, we analyzed the Akt/FoxO signalling pathway and its downstream targets Atrogin-1 and MuRF-1 via Western blotting. Dex treatment reduced Akt and FoxO3a phosphorylation levels, thereby increasing Atrogin-1 and MuRF-1 expression (Figure 3J). Notably, R-irisin administration effectively suppressed Atrogin-1/MuRF-1 upregulation while restoring Akt/FoxO3a phosphorylation in Dex-treated muscles. Strikingly, R-irisin further ameliorated Dex-induced atrophy by elevating myogenic regulatory factors (MyHC, Myogenin, and MyoD) and downregulating the muscle growth inhibitory factor MSTN (Figure 3I). Additionally, R-irisin targets the mTOR signaling pathway and the phosphorylation of its downstream effectors, P70S6K and 4E-BP1—a key regulatory node in the muscle anabolic pathway. (Figure 3J). Functional validation using puromycin incorporation assays in mature myotubes demonstrated that R-irisin significantly enhanced protein synthesis in both control and Dex-treated cells (Figure 3K). These findings collectively reveal that R-irisin counteracts Dex-induced muscle wasting through dual mechanisms: (1) inhibiting catabolic pathways (Atrogin-1/MuRF-1-mediated proteolysis) and (2) stimulating anabolic processes (protein synthesis via mTORC1 activation).

Muscle atrophy is associated with mitochondrial homeostasis loss, characterized by reduced mitochondrial density (volume), impaired biogenesis, and defective energy production [21]. R-irisin administration ameliorated these defects by enhancing mitochondrial ATP generation, citrate synthase activity, and aconitase activity in Dex-treated muscles (Figure 4A–C). Western blot analyses further confirmed that R-irisin upregulated mitochondrial biogenesis markers PGC-1α, COX2, and TOM20 in Dex-exposed myotubes (Figure 4D). Notably, R-irisin restored redox balance by reducing mitochondrial ROS levels (Figure 4E). These metabolic improvements coincided with structural remodelling of mitochondrial morphology, as evidenced by mitochondrial reduced fragmentation (Figure 4F). Collectively, these findings demonstrate that R-irisin counteracts Dex-induced mitochondrial damage through coordinated enhancement of mitochondrial morphology, biogenesis, and bioenergetic function.

SIRT1 plays a pivotal role in regulating skeletal muscle remodelling. To validate whether irisin directly interacts with SIRT1, we performed virtual docking analysis. The binding affinity of the irisin-SIRT1 complex was calculated to be −12.7 kcal/mol, indicating a strong binding interaction. The detailed interaction interfaces, including key residues involved in the binding, are illustrated in the figure (Figure 5A). Furthermore, treatment with 50 μM dexamethasone significantly reduced SIRT1 levels in C2C12 myotubes (Figure 5B). Strikingly, R-irisin rescued SIRT1 expression in a concentration-dependent manner (Figure 5B). Fluorometric assays further confirmed that R-irisin dose-dependently restored SIRT1 deacetylase activity (Figure 5C). In vivo, R-irisin administration reversed Dex-induced SIRT1 suppression in muscle tissues (Figure 5D). Mechanistically, Dex-mediated SIRT1 inhibition increased FoxO3a acetylation, driving upregulation of atrophy-related proteins atrogin-1, MuRF-1, and MSTN, while suppressing myogenic regulatory factor MyHC. R-irisin restored SIRT1 deacetylase activity, thereby suppressing FoxO3a hyperacetylation and downregulating atrogin-1/MuRF-1/MSTN expression (Figure 5E,F). Notably, co-treatment with the SIRT1-specific inhibitor EX-527 abolished R-irisin’s anti-atrophic effects (Figure 5E,F). Furthermore, R-irisin’s ability to rescue Dex-induced deficits in protein synthesis—assessed via puromycin incorporation assays—was dependent on SIRT1 activity (Figure 5G). These findings establish that R-irisin exerts its protective effects through SIRT1-mediated deacetylation pathways, coordinately regulating proteostasis and myogenesis.

PGC-1α, a key substrate of SIRT1 deacetylation and its biologically active form ¹³, was significantly hyperacetylated in Dex-treated C2C12 myotubes. Immunoprecipitation (IP) revealed that R-irisin treatment reversed Dex-induced elevation of acetylated PGC-1α (Figure 6A). Consistent with these findings, co-administration of the SIRT1 inhibitor EX-527 abolished R-irisin’s effects on both total and acetylated PGC-1α levels (Figure 6A). Western blot analyses further demonstrated that R-irisin-mediated restoration of mitochondrial biogenesis markers and function was blocked by EX-527 (Figure 6B). Mitochondrial membrane potential staining showed that Dex induced a collapse in membrane potential, which was rescued by R-irisin but reversed by EX-527 co-treatment (Figure 6C). Collectively, these results establish that R-irisin alleviates Dex-induced atrophy and mitochondrial dysfunction by restoring SIRT1-dependent deacetylation of FoxO3a and PGC-1α, thereby reactivating their pro-myogenic and bioenergetic functions.

Building on R-irisin’s robust in vitro efficacy, we extended our investigation to in vivo models. R-irisin administration effectively restored Dex-induced losses in gastrocnemius and tibialis anterior muscle mass, while co-administration with the SIRT1 inhibitor EX-527 antagonized these therapeutic effects (Figure 7A,B). Similarly, R-irisin-enhanced muscle functional performance-including grip strength, grid hanging capacity, and treadmill exhaustive distance-was nearly abolished by EX-527 co-treatment (Figure 7C–E). Histological analysis revealed that R-irisin prevented Dex-induced reductions in myofibre cross-sectional area (Figure 7F,G) and suppressed expression of key atrophy-related proteins: Atrogin-1, MuRF-1, and MSTN (Figure 7H), effects that were reversed by EX-527. Notably, R-irisin rescued Dex-mediated mitochondrial dysfunction, evidenced by restored mitochondrial integrity (Figure 8A), enhanced ATP concentration, citrate synthase activity, and aconitase activity (Figure 8B–D). These mitochondrial improvements coincided with upregulated biogenesis markers PGC-1α, COX2, and Tom20 (Figure 8E), all of which were antagonized by EX-527. Collectively, these in vivo findings confirm that R-irisin mitigates Dex-induced muscle atrophy and mitochondrial dysfunction through SIRT1-dependent pathways, reinforcing its therapeutic potential for sarcopenic disorders.